Mutational dissection of a hole hopping route in a lytic polysaccharide monooxygenase (LPMO)

Oxidoreductases have evolved tyrosine/tryptophan pathways that channel highly oxidizing holes away from the active site to avoid damage. Here we dissect such a pathway in a bacterial LPMO, member of a widespread family of C-H bond activating enzymes with outstanding industrial potential. We show that a strictly conserved tryptophan is critical for radical formation and hole transference and that holes traverse the protein to reach a tyrosine-histidine pair in the protein’s surface. Real-time monitoring of radical formation reveals a clear correlation between the efficiency of hole transference and enzyme performance under oxidative stress. Residues involved in this pathway vary considerably between natural LPMOs, which could reflect adaptation to different ecological niches. Importantly, we show that enzyme activity is increased in a variant with slower radical transference, providing experimental evidence for a previously postulated trade-off between activity and redox robustness.


Supplementary methods
Melting temperature SYPRO ® orange dye from Invitrogen was used to monitor the apparent melting temperature (Tm) of LPMOs 1 .Fluorescence by this dye is naturally quenched and is enhanced when it binds to hydrophobic residues of proteins that become exposed upon unfolding.A temperature ramp from 25 to 98 ºC with an increase of 1.5 ºC/min was used to evaluate the Tm.The SYPRO® orange stock solution (5000x) was diluted to an 8x stock in Milli-Q H2O prior to adding it for 1x working concentration in the reaction.The reactions contained 30 µM Cu 2+saturated LPMO and 1x SYPRO® Orange dye in 50 mM Tris pH 7.0.Reactions were prepared in quadruplicates, including control reactions without enzyme.
The fluorescence change was monitored with a StepOnePlus ™ Real-Time PCR (ThermoFisher Scientific).The StepOnePlus ™ software (v2.3) was used to obtain the negative first derivative of the normalized fluorescence signal with respect to temperature (-dF/dT), allowing an easy identification of the Tm as the lowest -dF/dT peak.

Substrate binding
Binding to β-chitin was evaluated under conditions similar to those used for LPMO activity measurements.Reactions were carried out at 22 ºC, with 1 mM ascorbate present to analyze the binding capacity of LPMO-Cu(I). 5 µM enzyme was added to mixtures of 10 g/L of β-chitin in 50 mM Tris, pH 8.0, with 1 mM ascorbate supplemented immediately before the addition of the LPMO.Samples were taken after 1, 3, 5, 10, 15, 30 and 60 minutes of incubation, and filtered using a 96-well filter plate (Merck) operated with a vacuum manifold (Millipore).
The unbound protein present in the soluble fraction was quantified with an adapted Bradford protocol for measuring low protein concentrations 2 .For each SmAA10A variant, a control reaction without β-chitin was included and used as reference to calculate the fraction (%) of unbound protein.Controls without enzyme were also added to monitor unspecific signals in the Bradford assay.All reactions were done in triplicates.

H2O2 Production Assay
H2O2 production was measured as previously described 3 .Stock solutions (10 mM) of Amplex TM Red Reagent (Thermo Fisher Scientific, Waltham, MA, USA) were prepared in DMSO.Reactions were prepared in a 90 μL volume containing 50 mM Tris, pH 8.0, 100 μM Amplex TM Red Reagent, 5 U/mL horseradish peroxidase (HRP, Sigma) and 2 μM LPMO, and pre-incubated at 30 °C for 5 minutes.Following pre-incubation, reactions were initiated with the addition of 10 μL of 10 mM ascorbate (1 mM final concentration) and the reaction mixtures were incubated at 30 °C.Formation of resorufin was monitored over 40 minutes at 540 nm in a Multiskan TM FC microplate photometer (Thermo Fisher Scientific, Waltham, MA, USA).A H2O2 standard curve was prepared in the same manner, with ascorbate added prior to the addition of Amplex TM Red Reagent and HRP 4 .

Product analysis by MALDI-TOF MS
Soluble products of β-chitin degradation by SmAA10A or its W119F variant were analyzed using a matrix-assisted laser desorption/ionization time-of-flight (MALDI-TOF) UltrafleXtreme mass spectrometer (Bruker Daltonics GmbH, Bremen, Germany). 1 μl of the filtered reaction sample was mixed with 2 μl of a matrix solution [9 mg/ml 2,5-dihydrooxybenzoic acid in 30% (v/v) acetonitrile] on the surface of an MTP 384-ground steel target plate (Bruker Daltonics).The target plate was air-dried, and MS data were collected using Bruker flexControl (version 3.4, build 169.5) software as described previously 5 .

Real-time monitoring of H2O2 turnover
The construction, use and experimental potential of the H2O2 sensor are described in detail in Schwaiger et al. 6 .The H2O2 sensor is based on the fast electrochemical detection of H2O2 using an Autolab potentiostat (PGSTAT101, Metrohm) connected to a rotating disk module (motor controller and rotating disk-setup, AUT.RDE.S, Metrohm).The measurements were performed using a three-electrode setup consisting of a gold rotating disk electrode (RDE, RDE.AU50.S, d = 5 mm, Metrohm) as the working electrode (WE), a coiled platinum wire (BASi) as the counter electrode (CE), and a Ag|AgCl electrode (3 M KCl, MF-2056, BASi) as the reference electrode (RE).The RE was protected from substrate poisoning by using a glass double junction (MF-2030, BASi) filled with 100 mM KCl.The stability of the reference electrode was checked on a regular basis by measuring the potential against the Labmaster electrode (EF-1352, BASi).The potential difference between the reference electrode and the Labmaster electrode is determined by measuring the open circuit potential, i.e. the equilibrium potential developed between the RE and the Labmaster electrode.To increase the sensitivity of the gold rotating disk working electrode, a thin layer of Prussian blue was deposited using cyclic voltammetry.The Prussian blue film is deposited on the surface of the electrode by cycling the WE eight to twelve times in a solution of 2.5 mM FeCl3, 2.5 mM K3[Fe(CN)6], 0.1 M KCl, and 0.1 M HCl in a potential window between 600-900 mV vs. SHE at a scan rate of 20 mV s -1 .After this deposition step, the WE was thoroughly rinsed with MilliQ-H2O and activated by electrochemical cycling in a solution of 0.1 M HCl and 0.1 M KCl between 160-590 mV vs. SHE at a scan rate of 50 mV s -1 .
The activated sensors were then rinsed with MilliQ-H2O, dried under a stream of N2, coated with 7 µL of Nafion (Merck, Darmstadt) and cured overnight at ambient atmosphere.On the next day, the prepared sensor was conditioned in the working buffer (50 mM Tris, pH 8.0 & 100 mM KCl) using the same conditions as used in the activation step.All measurements were performed using a water-jacketed low volume cell (MR-1212, BASi) connected to an SE-12 heating circulator (Julabo) to maintain a constant temperature of 37 °C.The H2O2 sensor was operated at an applied potential of 100 mV vs. SHE.
All experiments with the H2O2 sensor were performed with a volume of 4 mL and typically consisted of 3 essential steps: (i) The baseline was recorded until the signal (current, nA) of the H2O2 sensor was constant, then H2O2 was titrated into the electrochemical cell in 30 µM steps until a final concentration of 150 µM was reached.For each titration step, the signal was recorded for 30 s. (ii) LPMO was added to a final concentration of 1 µM and the reaction was recorded for another 30s.(iii) The reaction was started by the addition of 50 µM or 1 mM ascorbate and the time course of the reaction was either recorded until the baseline was reached or until the H2O2 time traces flattened out completely before reaching the baseline.The raw data obtained in current (nA) versus time (s) was converted to H2O2 concentration (µM) versus time (s) using the calibration function recorded at the start of every single experiment (derived from the initial stepwise titration of H2O2).Typically, the H2O2 sensors used in this study had sensitivities of 200-250 nA µM -1 cm -2 and a limit of quantification of 3-4 µM H2O2.Initial rates (µM s -1 ) of reaction were obtained by linear fitting to the initial part of the H2O2 depletion curves (usually the first 15 s).Only data based on fits with an R 2 of 0.98 or higher are reported.The data was acquired using the software NOVA 1.1 from Metrohm (Herisau, Switzerland).
Enzyme turnover numbers (TN, s -1 ) were obtained by subtracting the small but noticeable ascorbate-dependent depletion of H2O2 in the presence of 10 g L -1 βchitin from LPMO-catalyzed depletion.This background signal was routinely measured for each measurement series and ranges between 0.06 µM s -1 for 50 µM ascorbate and 0.19 µM s -1 for 1 mM ascorbate.The off-pathway peroxidaselike activity of the LPMO was determined in the absence of substrate, otherwise using the same conditions as defined above.Enzymatic turnover numbers were determined in triplicate, off-pathway peroxidase activity and background H2O2 depletion in duplicate.Supplementary Figure 5. Spectral subtractions for WT SmAA10A and its W178 variants.Spectral subtraction was performed on the spectra depicted in Fig. 2 of the main manuscript that show formation of features until maximum signal.There were nine or ten spectra, labeled from 1 (early; the first one) to 9 or 10 (late, the last one).The data were processed by subtracting the preceding spectrum from each of the spectra, as indicated in the Supplementary Figure 10.Oxidase activity of SmAA10A variants (2 µM), as determined with the Amplex Red assay performed in 50 mM Tris, pH 8.0, using 1 mM ascorbate as reductant.All initial rates (derived from linear progress curves for formation of resorufin) obtained are just above the ascorbate only control, indicating very slow H2O2 production for all variants tested.Note that H2O2 production by the LPMO is low compared to H2O2 production resulting from auto-oxidation of ascorbate and that he contribution of the LPMO will be even lower in reactions with an LPMO substrate, in which the LPMO oxidase activity will be surpressed 3,7,8  experiments, and no matter the background correction, the W119F variant consumed H2O2 faster than the WT.Note that these numbers imply that the background peroxidase activity of W119F was also consistently higher, compared to WT.Note also that the conditions in panel C are highly damaging, due to a very high H2O2 concentration, and that, while a reasonable linear fit was possible, the initial rates in fact are apparent initial rates with values that are severely affected by enzyme inactivation.
Inactivation of the LPMOs is evident from the fact that the baseline is not reached (i.e., no complete consumption of added H2O2).The curvatures in panels A-C show that, in all three cases, H2O2 consumption levels off faster for W119F, which shows that this variant is more readily damaged.Panel D incorporates an experiment identical to that shown in panel B (where the base line is reached) and shows what happens upon addition of another 150 M of H2O2.The much lower rates observed in this second reaction cycle show that considerable enzyme inactivation had taken place in the first reaction cycle, as would be expected when using these levels of H2O2.Note that, in this second cycle, the initial rate of the W119F variant no longer surpasses that of the wildtype, underpinning that this variant suffered more damage in the first cycle.While the wildtype maintained approximately 33 ± 7 % of its initial activity, the W119F variant maintained less than 10 %.
The perhaps somewhat surprising dependency of the catalytic rates on the concentration of ascorbate is a consequence of the extreme conditions: when H2O2 levels are high, the peroxidase reaction will be prominent, which leads to consumption of ascorbate and, eventually, enzyme inactivation 9 .

E-F:
Second order rate constants for reduction with ascorbate (E) and reoxidation with H2O2 (F) of WT SmAA10A and the variants W178Y, W178F and W119F in 50 mM Tris, pH 8.0, at 25 ºC, determined by linear regression analysis of rates determined at different concentrations of ascorbate and H2O2, respectively.Data are presented as means ± s.d.(n = 3).Panel E shows that the rates of ascorbate-driven reduction are similar for WT, W119F, W178Y and W178F, while the reoxidation rate is 3 to 4-fold higher for those enzymes containing the W178 in close proximity to the copper (WT and W119F).This indicates that W178 is involved in the (substrate-independent) turnover of H2O2.
55 % (LPMO-1) : 45 % (LPMO-2).Spectrometer conditions are described in the Methods.b: histidine brace in LPMOs showing the pH-dependent equilibrium between component LPMO-1 and LPMO-2.This agrees with previous observations for multiple AA10 LPMOs that each exhibit an EPR spectrum consisting of two species, with both a rhombic and an axial EPR component [11][12][13][14][15] .It was shown for several LPMOs, by EPR spectroscopy and computational studies, that the different EPR responses (components) are attributed to differences in the number of aquo/hydroxo ligands.For rhombic EPR spectra, a five-coordinate copper is expected, with two coordinating water species, while the axial EPR spectra are assigned to a four coordinate species.These water coordination differences have been studied by varying the sample's pH or binding of substrate [14][15][16] .
The 14 N superhyperfine pattern along in LPMO-2 is well reproduced by the simulation of the EPR with the inclusion of three strongly coupled nitrogens, A ~ 35 -45 MHz, confirming 3N-coordination environment of the histidine brace.
To investigate potential changes in the active site upon mutation, CW X-band EPR spectra of several variants (W119F, W178Y, W178M, W178F, W178Y/W119F) were measured, Supplementary Fig. 14.Overall, spectra with very similar responses were obtained for all variants.Most importantly, the 14 N superhyperfine pattern (~ 3,200 G -3,360 G), including both the number of lines and the magnitude of the splitting, is consistent among the samples.As described earlier for the simulation of LPMO-2, the 14 N hyperfine pattern is only reproduced by the inclusion of three strongly coupled nitrogens (i.e. two coordinating imidazoles and the N-terminus amine).The persistent nitrogen hyperfine pattern compared to the wild type enzyme confirms both the integrity of the Histidine-Brace and identical local copper coordination environments for the various variants, Supplementary Figs.14b and 14c.However, W178M exhibits a slightly larger line width, which precludes the observation of the 14 N superhyperfine splittings at high field positions.Nevertheless, g-values and the copper hyperfine remain constant, indicating no significant changes of the copper coordination sphere.
For the various variants studied, slight intensity shifts along can be observed, that are attributed to a change in the LPMO-1:LPMO-2 ratio of the two components.These changes may be the results of minor differences in sample pH, unknown second sphere effects that may influence water binding through altered hydrogen bonding networks and/or introducing additional strains (disorder) resulting in broadened spectra.Nevertheless, the influence of these minor changes is expected to be neglectable for the activity of the respective variants, particularly as the EPR spectra of subcomponent LPMO-2 offer a highresolution glimpse of the histidine brace and its uniformity in the presence of the distant aromatic mutations.

Supplementary Figure 2
. pH, reductant and temperature dependence of radical formation for SmAA10A-Cu(I) reacting with 20 molar equivalents of H2O2, detected in UV-vis stopped-flow absorption spectroscopy experiments.A-B: Bis-Tris, pH 6.5.C-D: sodium phosphate, pH 7.0.E-F: Tris, pH 8.0. 1 molar equivalent of ascorbate (A-D) or L-Cys (E-F) was used to generate LPMO-Cu(I) in situ.In panels A to F, the left panels show formation of signals, with the time until the maximum signal was observed indicated in the box.The right panels show the decay of such signals, with the time until full decay indicated in the box.The shapes of the spectra depicted in panels E and F (pH 8.0, L-Cys), as well as the derived rates of feature formation and decay, are very similar to those seen in experiments at pH 8.0 in which ascorbate was used to generate the reduced LPMO (Fig. 2 of the main manuscript).The experiments depicted in panels A to F were done at 4 ºC.Panel G illustrates the strong temperature dependence of the signals by showing spectra at maximum signal for the reaction shown for WT in Fig. 2 of the main manuscript, done at three temperatures.
Figure.Red color indicates early time points and blue color indicates later time points, with diminishing growth of the signals.The spectra have been vertically offset for better visual presentation of the data.The data for WT SmAA10A show features for both Trp• and Tyr• radicals are.The Trp• feature appears to saturate more quickly, leaving only a Tyr• radical signal in the difference spectra for the later time points.When W178 is mutated to Tyr in the W178Y containing variants, only a sharp, but broader, Tyr•-like feature is observed at a similar maximum wavelength than WT.In variants containing a Phe or a Met instead of Y178, a minute putative Y• feature became visible upon spectral subtraction.
. The bars represent the mean value ± s.d.(n = 3; independent measurements are shown as red dots).

Supplementary Table 1. Strategy to generate each variant gene and list of primers used in this study.
Experimental details can be found in the main manuscript's Methods section.